Secondary and Non-Target Adverse Effects

Environmental risk assessment must consider the unintended consequences of the environmental release of a transgenic plant, particularly as this may impact on existing agricultural practices and the agro-ecosystem. This discussion of potential unintended, or secondary, effects on non-target organisms is illustrated using examples that address the US Environmental Protection Agency’s (EPA’s) risk assessment methodology for determining adverse effects to non-target organisms. In the case of plant-pesticides, the intent of EPA’s approach is to evaluate the potential hazard to terrestrial wildlife, aquatic animals, plants and beneficial insects. If detrimental effects are observed under laboratory conditions, field studies are required to assess the actual abundance of non-target species under test and control conditions. In the field, insects, for example, are usually exposed to smaller amounts of toxin than the laboratory test dose because of diet choice and other environmental factors within the field setting. The choice of appropriate indicator organisms is based on the potential for field exposure to the novel protein expressed in transgenic plants, which is dependent on the tissue specificity of expression. For Bt crops where crop residue exposure is a possibility, EPA has required data on the toxicity of delta-endotoxins to birds (e.g., quail), fish, honeybees and certain other beneficial insects (e.g., ladybird beetles) and soil invertebrates (e.g., Collembola, earthworm species).

Non-Target Test Organisms

A non-target organism is any plant, animal or microorganism that is unintentionally impacted by the novel, or transgenic, plant. The following guidance on the selection of non-target test organisms has been adapted from US EPA data requirements for protein plant-pesticides:

1. Avian test species. Young bobwhite quail or mallard ducks between 14 and 28 days of age at the beginning of the test period.



2. Aquatic animals. This is relevant to Bt-expressing aquatic plants that may have applications in forests, drainage ditches, riverbanks, and partially aquatic crops such as rice. It also applies in the case of field crops that are grown near bodies of water.
   1. Freshwater fish species. EPA’s guidelines provide that the species tested be selected from the list of species recommended with the exception of goldfish (warmwater species--bluegill sunfish, channel catfish, and fathead minnow; coldwater species--rainbow trout, brook trout, coho salmon). These species are desirable test organisms for several important reasons: they are frequently used to evaluate chemical and microbial pesticides; EPA has considerable background data on these species; standard methods for the care and handling of these species are available; and the species are widely distributed, are generally available, and have a variety of food habits and habitat requirements.
      
      As appropriate, consideration should be given to testing species representative of the geographic region or ecosystem where the pesticidal plant will be cultivated. Fish species likely to scavenge intoxicated insects or the modified plant tissue (such as in farmed fish food) should be tested when appropriate. Unless there are other overriding considerations, the rainbow trout is recommended as the freshwater fish test species. It is a desirable test animal because it is partially insectivorous.
   
   
   
   2. Aquatic invertebrate species. The most likely plant tissue to be tested is pollen. Due to the broad phylogenetic spectrum from which the investigator may choose, it is difficult to select the most appropriate aquatic invertebrate. Daphnia, a Cladoceran, has the advantage of having considerable background data for comparative purposes. In addition, Daphnia exhibits a bioconcentration effect. This results from the filter feeding habits of Daphnia and is a desirable feature in terms of assuring that the test animal ingests the toxin containing tissue. Both Daphnia and certain aquatic insects have the advantage of a short life cycle and are useful for assessment of reproductive effects.
   
   
3. Non-target insect testing. Selection of the predator/parasite species to be tested should take into account such factors as the likelihood of exposure to the plant protein, phylogenetic proximity of the test species to target pest species, and similar relationships.

Assessment of potential non-target insect hazard is complicated by a number of factors. Many plant-pesticides are expected to be specifically chosen for their ability to control pest insects. In most cases, it can be assumed that the non-target insect group most at risk will be closely related to the pest species. While there are few non-target insects that have been shown to be economically important to humans, there are many non-target insects which have an important role in ecological processes and may benefit humans indirectly.

The host range is an important factor in hazard evaluation for a protein plant-pesticide. A problem here is that extrapolation, even across species lines, is often not dependable. For this reason, tests should be conducted with representatives from a number of “beneficial insect” taxa. EPA recommends that testing be performed on pollinator species, such as honey bee, and three other species of insects, representing at least two of the following groups—parasitic dipterans, predaceous hemipterans, predaceous coleopterans, predaceous mites, predaceous neuropterans, parasitic hymenopterans.

The requirements for evaluating the potential toxic effects of protein plant-pesticides on representative soil organisms, such as Collembola and earthworms, were originally based on the possibility of long-term exposure of these organisms to crop residues incorporated or left upon the soil surface. (The US EPA does not require such testing for registration of conventional pesticides or spray Bacillus thuringiensis products.) One of EPA’s reasons for requiring the non-target soil invertebrate tests was the concern that adverse effects on these species would cause a build up of plant detritus in cotton fields. EPA has since discovered that the long term soil use of highly toxic chemical insecticides, such as aldicarb, terbufos, phorate and carbofuran, which have long term effects on soil invertebrate species, has not resulted in the build-up of plant detritus in soils based upon available information on current routine agronomic practices. Some of these materials have half-lives of 10 or more years. Thus protein plant-pesticide crops, which are expected to have less impact on these species than the highly toxic chemical pesticides, should not result in any increased build up of plant detritus. Supporting this conclusion are data which indicate that Bt toxin production in plant-pesticides ceases at plant senescence in the majority of registered Bt maize crops, allowing some time for protein degradation prior to harvest. Additionally, the environmental fate data indicate that for currently registeredbtmaize crops only <1 to 90 grams of Bt protein per acre would enter the soil as a result of post harvest incorporation of Bt plants. Since proteins are known to degrade rapidly in the soil, the potential for significant soil buildup and hazard to non-target soil organisms is not anticipated from the growing of crops containing protein plant-pesticides.


The following discussion of the potential secondary effects of virus-resistant transgenic plants (e.g., phenotypic alterations and viral recombination) has been included as an example of another type of non-target effect.

Since the first report of transgenic plants expressing resistance to tobacco mosaic virus by Powell-Abel et al. in 1986, numerous transgenic crops tolerant or resistant to a wide range of viruses have been developed (Beachy 1997). These plants represent the first application of the concept of pathogen-derived resistance (Sanford & Johnston 1985) where viral genes encoding coat proteins (CP), replicases, defective movement proteins, proteases or helper components (Lomonossoff 1995) have been introduced into the plant genome. By far, the most common form of engineered pathogen-derived resistance is CP mediated resistance, which has been used to confer resistance to viruses in at least 13 RNA virus genera, including 23 distinct virus species (Grumet 1995).

Virus- and disease-resistant transgenic plants offer many potential agronomic and ecological benefits. This is clearly the case when no corresponding natural host resistance genes have been identified, or when it is not feasible to move these traits into commercial crops. There is also the potential to reduce the use of pesticides to control insect vectors of viral pathogens. Along with these benefits are potential ecological risks that should be considered. These risks, which relate to possible interactions between products of the viral transgene, either RNA or protein, and an infecting virus, include: synergism, heteroencapsidation, and recombination (Tepfer 1993; Robinson 1996; Aaziz & Tepfer 1999). Depending on the host plant, the consequences of movement of the virus resistance trait to wild relatives via outcrossing may also have to be considered.

Non-Persistent Phenotypic Effects
There are two concerns related to phenotypic alteration. The first is based on the possibility of an adverse synergistic interaction between the RNA or protein product of a transgene and an infecting heterologous virus (i.e., a virus other than the one used in construction of the transgenic plant, which also infects the host plant) such that there is an increase in the severity of the symptoms caused by infection with the heterologous virus. Synergistic interactions between different co-infecting viruses in a non-transgenic host plant have been described for a number of virus combinations. The first described and best-studied example is with potato virus X (PVX) and potato virus Y (PVY) (Rochow & Ross 1955). The underlying mechanism for these synergistic effects is not well known. Experimental systems have been devised to identify the responsible gene products by creating transgenic plants expressing various portions of the potyviral genome and examining symptom expression following inoculation with PVX. Based on these studies, the 5’-terminal 3544 nucleotides of the PVY genome, which encode protease-1 (P1), helper component-protease (HC-Pro) and protein-3 (P3), are essential for synergistic symptom expression (Vance et al. 1995).

The second concern is founded on the possibility of heteroencapsidation (also termed transencapsidation), in which the RNA (or DNA) genome of a heterologous infecting virus may be encapsidated by the transgene encoded coat protein. Heteroencapsidation could result in a one-time alteration in either the mode of transmission or host range of the invading heterologous virus. Heterologous encapsidation, also termed phenotypic mixing, is also possible under conditions of mixed infection with different viruses, as has been demonstrated for barely yellow dwarf luteovirus (Mathews 1991) and members of the potyvirus (Bourdin & Lecoq 1991) and tombusvirus (Dalmay et al. 1992) groups. Even though multiple virus infection is common in field grown plants and trees (Abdalla et al. 1985; Falk & Bruening 1994), field situations of transencapsidation have only been reported in a few instances involving insect transmission of viruses (Falk et al. 1995). Heterologous encapsidation has also been demonstrated in transgenic plants that express viral CP (Osburn et al., 1990; Dalmay et al., 1992; Holt & Beachy, 1992). For example, Lecoq et al. (1993) showed that when plants expressing a CP transgene derived from an aphid-transmissible strain of zucchini yellow mosaic potyvirus were challenged with a non-aphid transmissible strain (defective in CP not aphid transmission factor), a heterologous aphid transmissible strain was detected.

Both synergism and heteroencapsidation are short-term phenotypic effects that remain restricted to the vicinity of the transgenic crop, and will disappear without persistent effects on the environment if cultivation of the transgenic crop is halted. With respect to heterologous encapsidation, it “is not a problem, because it is limited to a single transfer, i.e., once a heterologously encapsidated genome is introduced into a new host, it reverts to using its own CP” (Henry et al. 1995). Generally, the effects of either synergy or heterologous encapsidation are not expected to be any more serious than the impacts that can occur in multiple virus infections of susceptible crops.

Viral Recombination
Viral RNA recombination has been proposed as the most significant risk associated with virus resistant transgenic plants. Before directly addressing this issue, it is important to examine the occurrence and significance of RNA recombination in mixed virus infections of non-transgenic plants. RNA recombination was first observed in co-infections of distinct strains of poliovirus carrying single mutations (Hirst 1962; Ledinko 1963), and since then has been documented for a growing number of RNA viruses. Both phylogenetic analyses of viral nucleotide sequences and laboratory experiments using non-replicative (animal viruses) or movement-defective (plant viruses) virus variants, in which the functional defect can be restored through RNA recombination, have been used as evidence for RNA recombination (Table 1). Genome replication within RNA viruses involves a virus-encoded RNA-dependent RNA polymerase (RdRp), and the most accepted model for viral RNA recombination is RdRp-mediated template switching mechanism. In this model, RNA recombination occurs during RNA synthesis if the RdRp pauses (on the donor strand) and switches to another site on the same template or to another strand (acceptor strand) to resume nascent RNA synthesis (Cooper et al. 1974; Nagy & Simon 1997).

It is nearly impossible to estimate the frequency of RNA recombination in mixed-infections of wild-type viruses. Although laboratory studies under conditions of high selective pressure have demonstrated that the activity of functionally defective mutant viruses can be restored through RNA recombination with the same, or closely related, wild-type viruses, there have been no published reports of RNA recombination between co-infecting wild-type viruses. RNA recombination requires that the two viruses undergo RNA replication at the same time in the same sub-cellular location, a set of conditions that can be difficult to achieve. This is illustrated by cucumber mosaic virus (CMV) and tomato aspermy virus (TAV), two members of the cucumovirus group, in which successful co-infection of tobacco plants is the exception, not the rule, with CMV generally excluding TAV (Stackey & Francki 1990).

In experimental systems, similar in concept to those developed to demonstrate RNA recombination between viruses, RNA recombination between a transgene transcript and an infecting virus RNA has been reported for cauliflower mosaic virus (CaMV), cowpea chlorotic mottle virus (CCMV), and tomato bushy stunt virus (TBSV) (Borja et al. 1999). For example, inoculation of transgenic Brassica napus expressing CaMV ORF VI, which encodes a movement protein necessary for systemic infection by the virus, with a movement protein defective strain of the virus resulted in the generation of recombinant CaMV particles (Gal et al. 1992). Similarly, Greene & Allison (1994, 1996) were able to demonstrate RNA recombination between a transgene encoding a 3’-terminal portion of the CCMV coat protein and an isolate of CCMV lacking this region of the coat protein gene.

Given that RNA recombination, both between viruses in mixed infections of non-transgenic plants, and between a transgene and a virus infecting a transgenic plant, can occur, the essential biosafety question is: will recombination between transgene mRNAs and infecting viruses lead to the creation of strains with novel and more damaging properties? To date, there is no empirical evidence supporting this contention. When isolates of recombinant CCMV were tested on a range of host plants, four out of seven displayed novel symptoms (Allison et al. 1997), but none of them was more fit than the parental strain when co-inoculated with wild-type virus (Allison et al. 1999). Other experiments examining the symptomatology of recombinant viruses, created in vitro by exchanging RNA segments between different strains or between different related viruses, also found occasional striking changes (Ding et al. 1996; Salanki et al. 1997; Carrere et al. 1999). However, these studies did not fully investigate the fitness of the recombinant viruses relative to the parental strains.

Although there is a scarcity of empirical evidence, there are no indications to date that the nature of viral recombinants arising from RNA recombination in mixed-virus infections of non-transgenic plants is different from those possible in singly infected transgenic plants. This being the case, it is reasonable to conclude that plants expressing viral sequences do not present new or significantly increased risks relative to what exists naturally in a “non-transgenic world”.
Plant pest potential
A plant may be considered a pest but not a weed. For example, a plant that produces an allelopathic substance may be considered a pest if the toxin produced has an undesirable environmental effect. Transgenic plants expressing novel toxins or potential allergens must be assessed accordingly.

There is no evidence to indicate that MON 810 has altered plant pest potential in comparison with its non-transformed counterpart.

Effects on Non-target organisms
There is extensive information on the lack of non-target effects from microbial preparations of Bacillus thuringiensis subsp. kurstaki (B.t.k.) containing the Cry1Ab protein. The full length Cry1Ab protein encoded by the cry1Ab gene used to produce MON 810, and the insecticidally active trypsin-resistant core protein (HD-1) produced in these plants, are identical to the respective full length and trypsin-resistant core Cry1Ab proteins contained in commercial microbial formulations that have been used safely for over 30 years (Lee et al., 1995a; EPA, 1988). B.t.k. proteins are extremely selective for the lepidopteran insects (MacIntosh et al. 1990; Klausner 1984; Aronson et al. 1986; Dulmage 1981; Whitely & Schnepf 1986), bind specifically to receptors on the mid-gut of lepidopteran insects (Wolfersberger et al. 1986; Hofmann et al. 1988a; Hofmann et al. 1988b; Van Rie et al. 1989; Van Rie et al. 1990) and have no deleterious effect on beneficial/non-target insects (Flexner et al. 1986; Krieg & Langenbruch 1981; Cantwell et al. 1972; EPA 1988; Vinson 1989; Mehn & Cozzi 1989).

Field Studies: United States
Field trials were conducted in the United States from 1993-95 to assess the impact of insect protected maize on beneficial arthropods. Maize inbreds and hybrids expressing the Cry1Ab protein were compared to their non-transformed counterparts for relative abundance of beneficial arthropods.

Methods:
1993 Trials: Four inbred maize lines were planted in one location in Iowa and one in Nebraska (8 rows/line/location): insect protected lines MON 801 and 523-06-1 and two non-transgenic inbreds. Non-destructive visual sampling was conducted to assess the numbers of key beneficial arthropods. Plants were inspected for the presence of the insects of interest. Counts were made on 5 plants/row on each sampling date.

1994 Trials: Four transformation events, 523-06-1, 546-09-1, MON 809 and MON 801, were evaluated at two locations in Iowa. A total of 12 lines were planted at the first location and 8 lines at the second with four replications per location. Sampling protocols were the same as those for 1993.

1995 Trials: Trial sites were established at two locations: Kentucky (6 insect-protected lines, 6 controls) and Iowa (MON 810 and control). Visual sampling was conducted three times.

Results:
1993: Adult O. insidiosus were the only predators in consistent abundance and so were the only species assessed. True replicates were not conducted and so no statistical analysis was conducted. The trend observed indicated that there were no adverse effects on predator numbers due to the presence of the Cry1Ab protein.

1994: Data are presented in Table 2. A statistically significant difference was observed for one line on one sampling date, when the number of predators was greater on the insect protected hybrid as compared to its non-transformed counterpart.



1995: The results from Kentucky are presented in Table 3 and from Iowa in Table 4. At the Kentucky site, three significant comparisons in the mean number of Orius adults were recorded but overall the numbers of Orius were comparable between transformed and non-transformed lines. In Iowa, differences were observed in the abundance of predators at each sampling time, but no differences were observed between insect-protected and non-transformed maize lines.






Conclusion:
The results observed in field studies from 1993-95 demonstrate that Cry1Ab-expressing maize had neither a direct nor an indirect effect on the beneficial arthropod species studied. Additionally, insect counts from Iowa in 1995 showed that insect-protected maize had no effect on spiders, coccinellid, chrysopid and nabids which are also known to be important predators of the European corn borer as well as of other economically important pests of maize.

Field Studies: France
In 1995 efficacy trials were established at two locations in France in order to evaluate the impact of MON 810 on populations of beneficial arthropods in comparison to its non-transformed counterpart (MON 810/Bt-) with or without insecticide.

Methods:
The trial was established as a full factorial design with presence/absence of each of three factors: Cry1Ab protein, ECB inoculation, and insecticide spray (delta-methrine at 20 g/ha). In addition to performance assessments, the number of beneficial arthropods was evaluated on 10 plants in each trial plot. In order to ensure the development of ECB infestation, half the plots were inoculated with the insect pest.

Results:
There were no significant differences in the beneficial arthropod populations when comparing the plots of MON 810 and un-treated MON 810/Bt- (Table 5). Significant differences in populations of beneficial arthropods were observed when comparing MON 810 with insecticide-treated MON 810/Bt- (Table 5).

Conclusion:
The Cry1Ab protein in MON 810 did not display any measurable adverse impact on populations of beneficial arthropods.

Laboratory Studies
Production of insecticidal proteins within plant matrices may prolong their environmental persistence and increase the bioavailability of the proteins to both target and non-target invertebrates (Jepson et al., 1994). The Cry1Ab trypsin-resistant core (HD-1) in ECB resistant maize is present in maize plant tissue remaining in the field after harvest. Post harvest maize plant residue is typically tilled into the soil but may also remain on the soil surface until the following year (no till). Results of a study conducted to assess the environmental fate (dissipation) of B.t.k. HD-1 protein in post harvest maize tissue determined that the B.t.k. HD-1 protein, as a component of post-harvest ECB resistant maize plants, will dissipate readily on the surface of, or cultivated into, soil (see Appendix 2).

Prior to eventual degradation, the Cry1Ab protein may be consumed by non-target soil invertebrates such as earthworms and Collembola, which are critical components of soil decomposition processes (Calow, 1993). Additionally, non-target insect toxicology tests are necessary for assessing potential ecological risks posed by exposure to novel crop varieties (Urban & Cook, 1986). Since B. thuringiensis insecticidal proteins are biologically active only after they have been ingested and bound to specialized receptors in the midgut of susceptible insects, dietary feeding tests are required to study the toxicological responses of non-target insect species.

As the Cry1Ab protein expressed in MON 810 is rapidly cleaved by digestive proteolytic enzymes following ingestion, its toxicological effects were assessed using bacterially expressed, trypsin digested, Cry1Ab trypsin resistant core protein (HD-1). The full-length native cry1Ab gene was introduced into Escherichia coli and expressed Cry1Ab protein was subjected to trypsin digestion, purified, characterized, and used for mammalian and beneficial insect safety assessment studies. The chemical and functional equivalence of the E. coli produced trypsin-resistant core HD-1 and Cry1Ab expressed in transgenic MON 810 plants was established using a rigorous set of criteria including molecular weight, immunoreactivity, and insecticidal activity (see Expressed Material / Effect).

The following are brief summaries of dietary feeding trials of key non-target indicator species. Details of each feeding trial, with the exception of parasitic hymenopteran and ladybird beetles, are presented in Appendix 3.

Honey bee larvae and adults:
These studies were performed to assess the safety of the Cry1Ab trypsin-resistant core protein for larvae and adult honey bee (Apis mellifera L.), a beneficial insect pollinator. A maximum hazard dose was used for these studies. The maximum nominal Cry1Ab protein concentration tested was greater than 10 times the estimated level required for 50% mortality (LC50) of several target pest Lepidoptera to the Cry1Ab protein (MacIntosh et al., 1990). No differences among treatments were observed and the LC50 for Cry1Ab protein in larval and adult honey bee was greater than 20 ppm, the highest dose tested. The no observed effect level was 20 ppm (Maggi & Sims, 1994a, 1994b; Sims, 1994).

Green lacewing:
Green lacewing (Chrysopa carnea) is a beneficial predatory insect commonly found in maize and other cultivated plants. There was no evidence that green lacewing larvae were adversely effected when fed moth eggs coated with a nominal concentration of 16.7 ppm Cry1Ab protein for seven days. Under the conditions of the test, the LC50 was greater than 16.7 ppm Cry1Ab protein, the highest dose tested (Hoxter & Lynn, 1992a).

Parasitic Hymenopteran:
A study was performed to assess the safety of the Cry1Ab trypsin-resistant core protein for parasitic Hymenopteran (Brachymeria intermedia), a beneficial parasite of the housefly (Musca domestics). Parasitic Hymenopteran exposed to trypsin-resistant Cry1Ab protein at a concentration of 20 ppm in honey/water solution for thirty days exhibited no treatment related mortality or signs of toxicity. The LC50 for Cry1Ab protein in parasitic Hymenopteran was greater than 20 ppm, the highest dose tested. The no-observed effect level was 20 ppm (Hoxter & Lynn, 1992b).

Ladybird beetles:
The ladybird beetle (Hippodamia convergent), is a beneficial predacious insect that feeds on aphids and other plant insects commonly found on stems and foliage of weeds and cultivated plants. Ladybird beetles exposed to trypsin-resistant Cry1Ab protein at a test concentration of 20 ppm in a honey/water solution for nine days exhibited no treatment related mortality or signs of toxicity. The LC50 for Cry1Ab protein in ladybird beetles was greater than 20 ppm, the highest dose tested. The no-observed effect level was 20 ppm (Hoxter & Lynn, 1992c).

Daphnia:
A study was performed to evaluate the acute toxicity of maize pollen containing the Cry1Ab protein to the cladoceran, Daphnia magna, during a 48-hour exposure period under static- renewal test conditions. Daphnids are representative of an important group of aquatic invertebrates. The estimated 48-hour LC50 value for D. magna exposed to CryIAb protein in maize pollen was >100 mg test pollen/l. There were no treatment-related effects observed at the 100 mg test pollen/l limit concentration.

Earthworm:
The objective of this study was to evaluate the toxicity of Cry1Ab insecticidal protein administered to earthworms during a 14-day exposure period in an artificial soil substrate. The 14-day LC50 value for earthworms exposed to Cry1Ab insecticidal protein was determined to be greater than 200 mg/kg dry soil, the single concentration tested. The no-observed-effect-concentration was 200 mg/kg dry soil.

Collembola:
The objective of this study was to examine the effect of plant-expressed Cry1Ab protein on Collembola, which are soil dwelling invertebrates that play a major role in soil ecosystems due to their feeding on decaying plant materials. Collembola could possibly be exposed to Cry1Ab protein in maize tissue remaining in fields after harvest. The results of this study indicate that, even at very high treatment levels, Collembola were not affected by chronic exposure to Cry1Ab in plants.

Northern Bobwhite Quail:
The purpose of this study was to assess the wholesomeness of ECB resistant maize meal from line MON 80187 when fed to quail (birds may feed on maize seed left in the field after harvest). No mortality occurred in birds fed up to 10% w/w (nominal 100,000 ppm) MON 80187 in the diet. The no-observed effect level was considered to be greater than 10% w/w. Based on the parameters measured, the wholesomeness of meal from ECB resistant maize line (MON 80187) was comparable to that of the control fine (MON 80087) when fed in the diet to quail.

1. Aaziz, R. & Tepfer, M. (1999). Recombination in RNA viruses and in virus-resistant transgenic plants. Journal of General Virology 80, 1339-1346.


2. Abdalla, O. A., Desjardins, P. R. & Dodds, J. A. (1985). Survey of pepper viruses in California by the ELISA technique. Phytopathology 75, 1311.


3. Allison, R.F., Thompson, C. & Ahlquist, P. (1990). Regeneration of a functional RNA virus genome by recombination between deletion mutants and requirement for cowpea chlorotic mottle virus 3a and coat genes for systemic infection. Proc. Natl. Acad. Sci. USA 87, 1820-1824.


4. Allison, R.F., Greene, A.E. & Schneider, W.L. (1997). Significance of RNA recombination in capsid-protein-mediated virus-resistant transgenic plants. In: “Virus-Resistant Transgenic Plants: Potential Ecological Impact”, pp. 40-44. Edited by M. Tepfer & E. Balazs. Versailles & Heidelberg: INRA & Springer-Verlag.


5. Allison, R.F., Schneider, W.L. & Deng, M. (1999). Risk assessment of virus-resistant transgenic plants. In: “Proceedings of the 5th International Symposium on Biosafety Results of Field Tests of Genetically Modified Plants and Microorganisms”, Braunschweig. Edited by J. Schiemann & R. Casper.


6. Aronson, A.I., Backman, W. & Dunn, P. (1986). Bacillus thuringiensis and related insect pathogens. Microbiol. Rev. 50, 1-24.


7. Beachy, R.N. (1997). Mechanisms and applications of pathogen-derived resistance in transgenic plants. Current Opinion in Biotechnology 8, 215-220.


8. Beck, D.L. & Dawson, W.O. (1990). Deletion of repeated sequences from tobacco mosaic virus mutants with two coat protein genes. Virology 177, 462-469.


9. Bergmann, M., Garcia-Sastre, A. & Palese, P. (1992). Transfection-mediated recombination of influenza A virus. Journal of Virology 66, 7576-7580.


10. Borja, M., Rubio, T., Scholthof, H.B. & Jackson, A.O. (1999). Restoration of wild-type virus by double recombination of tombusvirus mutants with a host transgene. Molecular Plant-Microbe Interactions 12, 153-162.


11. Bourdin, D. & Lecoq, H. (1991). Evidence that heteroencapsidation between two potyviruses is involved in aphid transmission of a nonaphid transmissible isolate from mixed infection. Phytopathology 28, 1459-1464.


12. Bujarski, J.J. & Kaesberg, P. (1986). Genetic recombination between RNA components of a multipartite plant virus. Nature 321, 528-531.


13. Calow, P. (ed.) (1993). Handbook of Ecotoxicology. Blackwell Scientific Publishing, London. 478 p.


14. Cantwell, G.E., Lehnert, T. & Fowler, J. 1972. Are biological insecticides harmful to the honey bees. Am. Bee J. 112, 294-296.


15. Carrere, I., Tepfer, M., Jacquemond, M. (1999). Recombinants of cucumber mosaic virus (CMV): determinants of host range and symptomatology. Arch Virol. 144(2), 365-79.


16. Cascone, P.J., Carpenter, C.D., Li, X.H. & Simon, A.E. (1990). Recombination between satellite RNAs of turnip crinkle virus. EMBO Journal 9, 1709-1715.


17. Cooper, P.D., Steiner-Pryor, A., Scotti, P.D. & Delong, D. (1974). On the nature of poliovirus genetic recombinants. Journal of Gneral Virology 23, 41-49.


18. Dalmay, T., Rubino, L., Burgyan, J., & Russo, M. (1992). Replication and movement of a coat protein mutant of cymbidium ringspot tombusvirus. Molecular Plant-Microbe Interactions 5, 379-383.


19. Ding, S.W., Shi, B.J., Li, W.X. & Symons R.H. (1996). An interspecies hybrid RNA virus is significantly more virulent than either parental virus. Proc Natl Acad Sci USA 93(15), 7470-4.


20. Dulmage, H.T. 1981. Microbial Control of Pests and Plant Diseases 1970-1980. (ed. Burges, H.D.) pp. 193-222. Academic Press, London.


21. Environmental Protection Agency (EPA). (1988). Guidance for the re-registration of pesticide products containing Bacillus thuringiensis as the active ingredient. NTIS PB 89-164198.


22. Falk, B. W. & Bruening, G. (1994). Will transgenic crops generate new viruses and new diseases. Science 263, 1395-1396.


23. Falk, B.W., Passmore, B.K., Watson, M.T. & Chin, L.-S. (1995). The specificity and significance of heterologous encapsidation of virus and virus-like RNAs. In: “Biotechnology and Plant Protection: Viral pathogenesis and disease resistance”. Bills, D.D. & Kung, S.-D. (eds.). World Scientific, Singapore, pp. 391-415.


24. Fernandez-Cuartero, B., Burgyan, J., Aranda, M.A., Salanki, K., Moriones, E. & Garcia-Arenal, F. (1994). Increase in the relative fitness of a plant virus RNA associated with its recombinant nature. Virology 203, 373-377.


25. Flexner, J.L., Lighthart, B. & Croft, B.A. (1986). The effects of microbial pesticides on non-target beneficial arthropods. Agric. Ecosys. Environ. 16, 203-254.


26. Gal, S., Pisan, B., Hohn, T., Grimsley, N. & Hohn, B. (1992). Agroinfection of transgenic plants leads to viable cauliflower mosaic virus by intermolecular recombination. Virology 187, 525-533.


27. Gal-On, A., Meiri, E., Raccah, B. & Gaba, V. (1998). Recombination of engineered defective RNA species produces infective potyvirus in planta. Journal of Virology 72, 5268-5270.


28. Galinat, W.C. (1988). The origin of corn. In Corn and Corn Improvement (Third Edition). eds. Sprague, G.F. and Dudley, J.W. pp. 1-31. American Society of Agronomy. Inc.., Crop Science Society of America Inc. and Soil Science Society of America. Inc. Madison, Wisconsin.


29. Greene, A.E. & Allison, R.F. (1994). Recombination between viral RNA and transgenic plant transcripts. Science 263, 1423-1425.


30. Greene, A.E. & Allison, R.F. (1996). Deletions in the 3’ untranslated region of cowpea chlorotic mottle virus transgene reduce recovery of recombinant viruses in transgenic plants. Virology 225, 231-234.


31. Grumet, R. (1995). Genetic engineering for crop virus resistance. HortScience 30, 449-456.


32. Harlow, E., & Lane, D. (1988). Immunoassay. In Antibodies: A Laboratory Manual. pp. 553-612.


33. Henry, C.M., Barker, I., Pratt, M., Pemberton, A.W., Farmer, M.J., Cotten, J., Ebbels, D., Coates, D. & Stratford, R. (1995). Risks associated with the use of genetically modified virus tolerant plants. A report to the Ministry of Agriculture Fisheries and Food (MAFF), United Kingdom.


34. Hirst, G.K. (1962). Genetic recombination with Newcastle disease virus, poliovirus and influencza. Cold Spring Harbor Symposia on Quantitative Biology 27, 303-308.


35. Hofmann, C., Vanderbruggen, H. V., Hofte, H., Van Rie, J. , Jansens, S. & Van Mellaert, H. (1988a). Specificity of B. thuringiensis delta-endotoxins is correlated with the presence of high affinity binding sites in the brush border membrane of target insect midguts. Proc. Natl. Acad. Sci. USA 85, 7844-7848.


36. Hofmann, C., Lüthy, P., Hutter, R. & Pliska, V. (1988b). Binding of the delta endotoxin from Bacillus thuringiensis to brush-border membrane vesicles of the cabbage butterfly (Pieris brassicae). Eur. J. Biochem. 173, 85-91.


37. Holt, C. A. & Beachy, R. N. (1992). In vivo complementation of infectious transcripts from mutant tobacco mosaic virus cDNA’s in transgenic plants. Virology 181, 109-117.


38. Hoxter, K.A. & Lynn, S.P. (1992a). Activated Btk HD-1 protein: a dietary toxicity study with green lacewing larvae. Study Number WL-92- 155, an unpublished study conducted by Monsanto Company and Wildlife International Ltd. EPA MRID no. 43468003.


39. Hoxter, K.A. & Lynn, S.P. (1992b). Activated Btk HD-1 protein: a dietary toxicity study with parasitic Hymenopteran (Brachymeria intemedia). Study Number WL-92-157, an unpublished study conducted by Monsanto Company and Wildlife International Ltd. EPA MRID no. 43468004.


40. Hoxter, K.A. & Lynn, S.P. (1992c). Activated Btk HD-1 protein: a dietary toxicity study with ladybird Beetles. Study Number WL-92-156, an unpublished study conducted by Monsanto Company and Wildlife International Ltd. EPA MRID no. 43468005.


41. Jepson, P.C., Croft, B.A., & Pratt, G.E. (1994). Test systems to determine the ecological risks posed by toxin release from Bacillus thuringiensis genes in crop plants. Molecular Ecology 3, 81-89.


42. Klausner, A. (1984). Microbial insect control. Bio/Technology 2, 408-419.


43. Kottier, S.A., Cavanagh, D. & Britton, P. (1995). Experimental evidence of recombination in coronavirus infectious bronchitis virus. Virology 213, 569-580.


44. Krieg, A. & Langenbruch, G.A. (1981). Susceptibility of arthropod species to Bacillus thuringiensis. In Microbial Control of Pests and Plant Diseases, (ed. Burges, H.D.) pp. 837-896. Academic Press, London.


45. Lai, M.M., Baric, R.S., Makino, S., Keck, J.G., Egbert, J., Leibowitz, J.L. & Stohlman, S.A. (1985). Recombination between nonsegmented RNA genomes of murine coronaviruses. Journal of Virology 56, 449-456.


46. Lee, T-C., Bailey, M., & Sanders, P.R. (1995). Compositional comparison of Bacillus thuringiensis subsp. kurstaki HD-1 protein produced in ECB resistant corn and the commercial microbial product, DIPEL. Study Number 94-01-39-12. an unpublished study conducted by Monsanto Company. EPA MRID no. 43533203.


47. Li, Y. & Ball, L.A. (1993). Nonhomologous RNA recombination during negative strand synthesis of flock house virus RNA. Journal of Virology 67, 3854-3860.


48. Lecoq, H., Ravelonandr, M., Wipf-Scheibel, Monision, M., Raccah, B. & Dunez, J. (1993). Aphid transmission of a non-aphid transmissible strain of zucchini yellow mosaic potyvirus from transgenic plants expressing the capsid protein of plum poxvirus. Molecular Plant-Microbe Interactions 3, 301-307.


49. Ledinko, N. (1963). Genetic recombination with poliovirus type1: studies of crosses between a normal horse serum-resistant mutant and several guanidine-resistant mutants of the same strain. Virology 20, 107-119.


50. Lomonossof, G.P. (1995). Pathogen-derived resistance to plant viruses. Annual Review of Phytopathology 33, 323-343.


51. MacIntosh, S.C., Stone, T.B. Sims, S.R., Hunst, P., Greenplate,J.T., Marrone, P.G., Perlak, F.J., Fischhoff, D.A.& Fuchs, R.L. (1990). Specificity and efficacy of purified Bacillus thuringiensis proteins against agronomically important insects. J. Insect Path. 56, 258-266.


52. Maggi, V.L. & Sims, S.R. (1994a). Evaluation of the dietary effects of purified B.t.k. endotoxin protein on honey bee larvae. Study Number IRC-91-ANA-13, an unpublished study conducted by Monsanto Company and California Agricultural Research, Inc. EPA MRID no. 43439202.


53. Maggi, V.L. & Sims, S.R.. (1994b). Evaluation of the dietary effects of purified B.t.k. endotoxin proteins on honey bee adults. Study Number IRC-91-ANA-12. an unpublished study conducted by Monsanto Company and California Agricultura lResearch, Inc. EPA MWD no. 43439203.


54. Matthews, R. E. F. (1991). Plant Virology. Academic Press, New York. 835pp.


55. Matsudaira, P. (1987). Sequence from picomole quantities of proteins electro blotted onto polyvinylidene difluoride membranes. J. Biol. Chem. 262, 10035-10038.


56. McCahon, D., King, M.Q., Roe, D.S., Slade, W.R., Newman, J.W.I. & Cleary, A.M. (1985). Isolation and biochemical characterization of intertypic recombinants of foot-and-mouth disease virus. Virus Research 3, 87-100.


57. Melin. B.E. & Cozzi, E.M. (1989). In Safety of Microbial Insecticides (eds. Laird, M., Lacey. L.A. and Davidson, E.W.) pp. 150-167. CRC Press, Boca Raton, FL.


58. Munishkin, A.V., Veronin, L.A. & Chetverin, A.B. (1988). An in vivo recombinant RNA capable of autocatalytic synthesis by Qbeta replicase. Nature 333, 473-475.


59. Nagy, P.D. & Simon, A.E. (1997). New insights into the mechanisms of RNA recombination. Virology 235, 1-9.


60. Onodera, S., Qiano, X., Gottlieb, P., Strassman, J., Frilander, M. & Mindish, L. (1993). RNA structure and heterologous recombination in the double-stranded RNA bacteriophage phi6. Journal of Virology 67, 4914-4922.


61. Osburn, J. K., Sarkar, S. & Wilson, T. M. A. (1990). Complementation of coat protein-defective TMV mutants in transgenic plants expressing coat protein. Virology 179, 921-925.


62. Powell-Abel, P., Nelson, R.S., De, G., Hoffmann, N., Rogers, S.G., Fraley, R.T. & Beach, R.N. (1986). Delay of disease development in transgenic plants that express the tobacco mosaic virus coat protein gene. Science 232, 738-763.


63. Robinson, D.J. (1996). Environmental risk assessment of releases of transgenic plants containing virus-derived inserts. Transgenic Research 5, 359-562.


64. Rochow, W.F. & Ross, A.F. (1955). Plant Disease (Reporter) 52, 344-358.


65. Salanki, K., Carrere, I., Jacquemond, M., Balazs, E. & Tepfer, M. (1997). Biological properties of pseudorecombinant and recombinant strains created with cucumber mosaic virus and tomato aspermy virus. J Virol. 71(5), 3597-602.


66. Sanders, P.R., Elswick, E.N., Groth, M.E. & Ledesma, B.E. (1995). Evaluation of insect protected corn lines in 1994 US field test locations. Study Number 94-01-39-01, MSL-14179, an unpublished study conducted by Monsanto Company. EPA MRID no. 43665502.


67. Sanford, J.C. & Johnston, S.A. (1985). The concept of parasite-derived resistance: deriving resistance genes from the parasite’s own genome. Journal of Theoretical Biology 113, 395-405.


68. Sims, S.R. (1994). Stability of the CryIA(b) insecticidal protein of Bacillus thuringiensis var. kurstaki (B.t.k. HD-1) in sucrose and honey solutions under non-refrigerated temperature conditions. Study Number IRC-91-ANA-11, an unpublished study conducted by Monsanto Company. EPA MRID no. 43468002.


69. Stackey, S.T. & Francki, R.I.B. (1990). Interaction of cucumoviruses in plants: persistence of mixed infections of cucumber mosaic and tomato aspermy viruses. Physiological and Molecular Plant Pathology 36, 409-419.


70. Tepfer, M. (1993). Viral genes and transgenic plants: what are the potential environmental risks? Bio/Technology 11, 1125-1132.


71. Urban, D.J. & Cook, N.J. (1986). Standard Evaluation Procedure for Ecological Risk Assessment. EPA/540/09-86/167, Hazard Evaluation Division, Office of Pesticide Programs, US Environmental Protection Agency, Washington D.C.


72. Vance, V.B., Berger, P.H., Carrington, J.C., Hunt, A.G. & Shi, X.M. (1995). 5' proximal potyviral sequences mediate potato virus X/potyviral synergistic disease in transgenic tobacco. Virology 206, 583-590.


73. van der Kuyl, A.C., Neeleman, L. & Bol, J.F. (1991). Complementation and recombination between alfalfa mosaic virus RNA3 mutants in tobacco plants. Virology 183, 731-738.


74. Van Rie, J., Jansens, S. Hofte, H., Degheele, D. & Van Mellaert, H. (1989). Specificity of Bacillus thuringiensis delta-endotoxins, importance of specific receptors on the brush border membrane of the mid-gut of target insects. Eur. J. Biochem. 186, 239-247.


75. Van Rie. J., Jansens, S., Hofte, H., Degheele, D. & Van Mellaert, H. (1990). Receptors on the brush border membrane of the insect midgut as determinants of the specificity of Bacillus thuringiensis delta-endotoxins. Appl. Environ. Microbiol. 56, 1378-1385.


76. Vinson. S.B. (1989). Potential impact of microbial insecticides on beneficial arthropods in the terrestrial environment. In Safety of Microbial Insecticides. (eds. Laird, M., Lacey, L.A. and Davidson, E.W.) pp. 43-64. CRC Press, Inc. Boca Raton, FL.


77. Weiss, B.G. & Schlesinger, S. (1991). Recombination between Sindbis virus RNAs. Journal of Virology 65, 4017-4025.


78. White, K.A. & Morris, T.J. (1994). Recombination between defective tombusvirus RNAs generates functional hybrid genomes. Proc. Natl. Acad. Sci. USA 91, 3642-3646.


79. Whitely, H.R. & Schnepf, H.E. (1986). The molecular biology of parasporal crystal body formation in Bacillus thuringiensis. Ann. Rev. Microbiol. 40, 549-576.


80. Wolfersberger, M.G., Hofmann, C. & Luthy, P. (1986). Interaction of Bacillus thuringiensis delta-endotoxin with membrane vesicles isolated from lepidopteran larval midgut. In bacterial protein toxins. eds. Falmagne, P., Fehrenbech, F.J., Jeljaszewics, J. and Thelestam, M.. Gustav Fischer, pp. 237-238. New York, New York, U.S.A.